Báo cáo khoa học: The pivotal regulator GlnB of Escherichia coli is engaged in subtle and context-dependent control potx

The pivotal regulator GlnB of Escherichia coli is engaged in subtle and context-dependent control Wally C. van Heeswijk 1 , Douwe Molenaar 1 , Sjouke Hoving 1, * and Hans V. Westerhoff 1,2 1 Department of Molecular Cell Physiology, Faculty of Earth and Life Sciences, Vrije Universiteit, Amsterdam, The Netherlands 2 Manchester Centre for Integrative Systems Biology, University of Manchester, UK Because the environment changes frequently for many unicellular organisms, subtle regulation may be impor- tant for relative fitness. Appropriate adaptation requires a precise response to an accurate assessment of environmental changes. In some systems, the signal is transduced by the reversible covalent modification of a protein cascade, without transferring a chemical group down the chain. The functional activity of the protein at the bottom of the hierarchy depends on the modification state of that protein. The advantage of modulating the activity of a protein by a cascade-type of regulation rather than by allosteric interaction remains unclear. Based on theoretical analysis, it has been argued that regulatory cascades might serve the function of high signal amplification [1–9]. Here, we suggest that the opposite may be the case: they may serve the function of subtlety of regulation. In stark contrast to the number of theoretical sug- gestions, little is known experimentally about the extent to which the various proteins participating in a Keywords glutamine synthetase; metabolic control analysis; P II; signal transduction cascades; ultrasensitivity Correspondence W. C. van Heeswijk, Faculty of Earth and Life Sciences, Department of Molecular Cell Physiology, Vrije Universiteit, De Boelelaan 1085, NL -1081 HV Amsterdam, The Netherlands Fax: +31 20 598 7229 Tel: +31 20 598 7228 E-mail: wc_van_heeswijk@hotmail.com Website: http://www.bio.vu.nl/vakgroepen/ mcp/main/index.html *Present address Novartis Institutes of Biomedical Research, Basel, Switzerland (Received 5 February 2009, revised 3 April 2009, accepted 8 April 2009) doi:10.1111/j.1742-4658.2009.07058.x This study tests the purported signal amplification capability of the gluta- mine synthetase (GS) regulatory cascade in Escherichia coli. Intracellular concentrations of the pivotal regulatory protein GlnB were modulated by varying expression of its gene (glnB). Neither glnB expression nor P II * (i.e. the sum of the concentration of the P II -like proteins GlnB and GlnK) had control over the steady-state adenylylation level of GS when cells were grown in the presence of ammonia, in which glnK is not activated. Follow- ing the removal of ammonia, the response coefficient of the transient deadenylylation rate of GS–AMP was again zero with respect to both glnB expression and P II * concentration. This was at wild-type P II * levels. A 20% decrease in the P II * level resulted in the response coefficients increasing to 1, which was quite significant yet far from expected for zero-order ultrasensi- tivity. The transient deadenylylation rate of GS–AMP after brief incubation with ammonia was also measured in cells grown in the absence of ammonia. Here, GlnK was present and both glnB expression and P II * lacked control throughout. Because at wild-type levels of P II *, the molar ratio of P II *-tri- mer ⁄ adenylyltransferase-monomer was only slightly above 1, it is suggested that the absence of control by P II * is caused by saturation of adenylyltrans- ferase by P II *. The difference in the control of deadenylylation by P II * under the two different growth conditions indicates that control of signal transduction is adjusted to the growth conditions of the cell. Adjustment of regulation rather than ultrasensitivity may be the function of signal trans- duction chains such as the GS cascade. We discuss how the subtle interplay between GlnB, its homologue GlnK and the adenylyltransferase may be responsible for the ‘redundant’, but quantitative, phenotype of GlnB. Abbreviations ATase, adenylyltransferase; GS, glutamine synthetase; IPTG, isopropyl b- D-1-thiogalactoside; MCA, metabolic control analysis; UTase, uridylyltransferase. 3324 FEBS Journal 276 (2009) 3324–3340 ª 2009 The Authors Journal compilation ª 2009 FEBS regulatory cascade control signal transduction in vivo. Statements like ‘this protein is or is not involved’ do not suffice if subtlety of regulation is the issue. The relative extents to which the various proteins control signal transduction in the physiological state needs to be addressed. Does a (small) change in the activity of one of these proteins affect the strength of the response to the signal and does such a change interfere with the rate of signal transfer through the chain? Because phenotypes may be quantitative and subtle, analysis of knockout strains may not suffice. To address these questions a method is needed to quantify the magnitude of the control exercised by a protein on a physiological function, as well as the extent to which that magnitude changes with the conditions. Such methods have been developed for the control by enzymes of the fluxes through metabolic pathways. One of these is known as metabolic control analysis (MCA) [10–12]. In this method, the activity of the rele- vant enzyme (e i ) is modulated by inhibitor titration [13] or gene-expression titration [14,15] and the relative effect on the physiological property of interest, e.g. flux (J) through the pathway, is measured to give the flux–control coefficient (C J ei ) (i.e. the intrinsic control of the modulated enzyme on the flux). The activity of some enzymes can be modulated by the binding of an allosteric effector, e.g. a regulatory protein (A). If one modulates the concentration of the latter, then the relative effect on the physiological property of interest, e.g. flux through the pathway (J), divided by the rela- tive (small) modulation of the effector concentration gives the flux–response coefficient (R J A ), whereas the effect of A on the local rate (v ei ) of enzyme e i is quan- tified by an elasticity coefficient (e m ei A ) [10]. The response, control and elasticity coefficients relate to each other through R J A ¼ C J ei Á e m ei A [10]. In signal trans- duction cascades, the steady-state response of a steady fraction of a modified enzyme to an effector molecule may be called signal amplification if the corresponding response coefficient is > 1 [16]. Here, we use the MCA approach both conceptually and experimentally to address the question of how intensely a regulatory protein controls signal transduc- tion. Because the glutamine synthetase (GS) adenylyla- tion cascade has been studied extensively at the genetic and molecular (e.g. kinetic) levels [17,18], we used this cascade as the experimental model system; GS catalyses the incorporation of ammonia into gluta- mate to form glutamine [19]. Glutamine is a precursor at a branch point for several biosynthetic pathways [20]. GS is a homo-dodecameric protein [21]. This key enzyme in nitrogen anabolism is regulated at three levels: allosteric regulation, post-translational modifi- cation and transcriptional regulation [17–19]. The covalent modification of GS is regulated by a dual, bicyclic cascade (Fig. 1). GS can be both adenylylated and deadenylylated by the bifunctional enzyme ade- nylyltransferase (ATase) [22]; the N-terminal domain of ATase carries the deadenylylation activity, the C-terminal domain carries the adenylylation activity [23–25]. Covalent modification of all 12 subunits of GS (GS 12 ) to yield GS 12 –AMP 12 results in an almost inactive enzyme. Adenylylation of GS is stimulated by the protein GlnB, whereas deadenylylation is stimu- lated by the modified GlnB (GlnB–UMP) protein [22]. N-poor GlnK 3 GlnK 3 –UMP 1–3 GlnB 3 –UMP 1–3 UTase N-poor N-rich UTase GlnB 3 N-rich GS 12 –AMP 1–12 + + GS 12 ATase + + glu + NH 3 gln Fig. 1. The GS adenylylation dual bicyclic cascade in Escherichia coli. The activity of GS which catalyses the incorporation of ammonia (NH 3 ) into glutamate (glu) forming glutamine (gln), is regulated by a dual bicyclic cascade. Only the protein components are shown; addi- tional substrates and products and the small molecule effectors, glu- tamine and 2-oxoglutarate, of the four reactions are not included. Reactions catalysed by the bifunctional enzymes UTase (EC 2.7.7.59) and ATase (EC 2.7.7.49) are shown as solid curved arrows. Details and kinetics of the reactions catalysed by UTase and ATase have been described previously [22,26]. Stimulation of GlnB 3 , GlnB 3 – UMP 1–3 , GlnK 3 and GlnK 3 –UMP 1–3 are shown by thin right-angled arrows. +, stimulation. When cells are grown in N-poor medium (e.g. in the absence of ammonia but in the presence of glutamine), UTase catalyses the uridylylation of GlnB 3 and GlnK 3 forming GlnB 3 –UMP 1–3 and GlnK 3 –UMP 1–3 , respectively. The latter two stimulate ATase to deadenylylate GS 12 –AMP 1–12 into native and active GS 12 . Reversibly, when cells are grown in N-rich medium (e.g. in the presence of ammonia) or in the absence of ammonia and pulsed with ammonia, UTase catalyses the de-uridylylation of GlnB 3 –UMP 1–3 and GlnK 3 – UMP 1–3 forming native GlnB 3 and GlnK 3 , respectively. GlnB 3 and GlnK 3 stimulate ATase to adenylylate GS 12 into the almost inactive GS 12 –AMP 12 . However, in N-rich medium the expression of glnK is not activated [32,33,36] and therefore, in N-rich conditions GS is regulated by only one bicyclic cascade. W. C. van Heeswijk et al. GlnB: ultrasensitive versus subtle control FEBS Journal 276 (2009) 3324–3340 ª 2009 The Authors Journal compilation ª 2009 FEBS 3325 Modification of GlnB is catalysed by the bifunctional enzyme uridylyltransferase (UTase) [26]. GlnB is a homotrimeric protein [27,28] and all three subunits can be uridylylated. UTase may monitor the glutamine concentration and GlnB may monitor 2-oxoglutarate [26,29]. GlnB is also involved in the transcriptional regulation of glnA, the gene encoding GS, via the two-component regulation system NRI ⁄ NRII (NtrC ⁄ NtrB) (not shown in Fig. 1) [30,31]. GlnK, a para- logue of GlnB [32–34], is also a homotrimer [35], and can also stimulate the adenylylation reaction in vitro [33,34], however, it is less potent than GlnB [34]. In the presence of purified UTase or in extracts contain- ing overproduced UTase, GlnK can be modified to GlnK–UMP [33,34]. In N-poor media, glnK is expressed and GS is regulated by a dual bicyclic cascade (Fig. 1). In N-rich media, transcription of glnK is not activated [32,33,36] and GS should be regulated by only one bicyclic cascade. In this study, we focus on the deadenylylation reaction. To investigate the actual in vivo importance of GlnB, the cascade must be studied in a wild-type chromosomal background. Although helpful, a deletion strain missing one of the proteins operating in the cascade will not yield definitive information about the physiological state because its signal flux is completely interrupted. Delet- ing a parallel signal transduction pathway, e.g. by delet- ing the glnK gene, will artificially force signal transduction into the other route. Indeed, it has been shown for some growth conditions of Escherichia coli that GlnK can complement the absence of GlnB [33]. Borrowing a strategy from MCA, we therefore implemented a small modulation of the GlnB concen- tration in an otherwise wild-type environment, which should not, therefore, affect the regulation structure of the GS adenylylation system. We observed that the pivotal regulatory protein GlnB does not control the steady-state activity of GS. Its control of the deade- nylylation rate of GS–AMP depends on the growth history of the cells, but does not purport to signal amplification. Functional implications and the mecha- nistic basis for this conditional redundancy of GlnB (and GlnK) are discussed. Results Modulation of the GlnB concentration in vivo and the levels of GlnB and GlnK In order to modulate cellular GlnB activity around wild-type levels, we inserted a promoter cassette containing a lacI q1 gene and an isopropyl b-d-1-thio- galactoside (IPTG)-inducible, P A1lacO-1 promoter [37], upstream of the glnB gene at the wild-type chromo- somal location (Fig. 2). The P A1lacO-1 promoter–opera- tor sequence consisted of promoter P A1 of phage T7 combined with two lacO-1 operators, as constructed A * - P NotI* NotI glnBorfXB cam trpA term . lacI q1 A1lacO-1 RBS EcoNI* EcoNI* B 0 50 100 150 200 0 50 100 150 200 250 300 350 [PII* ng·mg –1 protein] [IPTG] (µ M ) Fig. 2. Modulation of the glnB expression by IPTG. (A) The IPTG- inducible promoter upstream of the glnB gene at the wild-type chromosomal location of strain WCH15. The drawing is not to scale. The promoter cassette was inserted as a NotI fragment into the EcoNI site upstream of the translation start of the glnB gene (NotI* and EcoNI* are blunted sites). The promoter cassette con- tains a cam gene for chloramphenicol-resistance, a synthetic trpA- transcriptional terminator [60], a LacI q1 gene [38,39] and a synthetic P A1LacO-1 promoter [37]. The ribosomal binding site (RBS, black box) is wild-type. Solid arrows indicate the orientation of transcription. The dotted arrow indicates the transcription start point of the IPTG- inducible promoter. (B) Intracellular P II * concentration as a function of the extracellular IPTG concentration (l M). Cells were grown in the presence of ammonia. Cultures are the same as in Fig. 3. [P II *] was measured by western blot analysis using polyclonal GlnB anti- body, as described in Materials and methods. For the [P II *] dataset (including error bars) see Fig. 3B. The error bars of the [P II *] values <25ngÆmg )1 protein are smaller than the symbol. Although this antibody cross-reacts with GlnK, [P II *] may regarded as being [GlnB] because glnK is not expressed in this medium. Closed cir- cles depict WCH15 grown in the presence of the indicated concen- tration of IPTG; the black line is a result of a linear regression calculation of the data points from 0 to 150 l M IPTG. Open circle, YMC10 (wild-type); open square, RB9060 (4glnB). The IPTG con- centration that should correspond with [P II *] of YMC10 and RB9060 was calculated by interpolation of the two most proximate [IPTG, P II *] data points of each strain. GlnB: ultrasensitive versus subtle control W. C. van Heeswijk et al. 3326 FEBS Journal 276 (2009) 3324–3340 ª 2009 The Authors Journal compilation ª 2009 FEBS by Lutz & Bujard [37]. The lacI q1 gene contained a promoter up-mutation which produced 100 times more repressor than wild-type cells [38,39]. The inducible promoter was inserted upstream of the glnB gene in a lacY deletion mutant (lacU169) in order to enhance the controllability of expression of the glnB gene by IPTG [40]. The combination of these three elements in strain WCH15 enabled us to titrate, using IPTG, the cellular GlnB concentration around wild-type levels (Fig. 2B). In the experiment shown in Fig. 2B, cells were grown in the presence of ammonia and the indi- cated IPTG concentration. The cellular GlnB concen- tration in the various cultures was analysed by western blot using a polyclonal GlnB antibody, as described in Materials and methods. Because the polyclonal GlnB antibody cross-reacted with paralogue GlnK, which has the same electrophoretic mobility as GlnB [32], the IPTG-dependent increase in the intensity of the GlnB band was quantified as the sum of the concentrations of GlnB and GlnK and was denoted by P II *. Note that P II * includes the modified forms of GlnB and GlnK as well, i.e. GlnB–UMP and GlnK–UMP. Because tran- scription of glnK is not activated in medium containing ammonia, [P II *] may be regarded as being [GlnB]. As shown in Fig. 2B, [P II *], and hence [GlnB], in the wild-type strain (YMC10) is 87 ngÆmg )1 protein. [P II *] of the DglnB strain and of WCH15 without IPTG is not completely zero because glnK may have some residual activity. The (minor) difference between the two strains may be because of inaccuracies in the western blot method. In cells grown in the presence of ammonia, GlnB at wild-type levels does not control the GS–AMP deadenylylation rate, although it does at lower levels WCH15 cells were grown overnight to A 600 $ 0.3 at various IPTG concentrations in minimal medium con- taining 22 mm glucose, 14 mm ammonia and 14 mm l-glutamine (N-rich). At this growth stage, almost all GS subunits were adenylylated (Fig. 3A; t = 0). The adenylylation state of GS was expressed in terms of the average number AMP moieties per GS dodecamer (n), as inferred from an activity assay (see Materials and methods). When we removed the ammonia plus glutamine from the medium, by pipetting the washed cells into the same medium without a nitrogen source [32,33] (see Materials and methods), we found that GS–AMP was de-modified towards GS, presumably because of a shift in the P II * ⁄ P II *–UMP ratio towards P II *–UMP. At various times after removal of the nitrogen source, the maximum deadenylylation rate, i.e. the rate in the inflection point of the curve in Fig. 3A, was calculated as described in Materials and methods. The cellular P II * concentration of the differ- ent cultures was measured by western blot analysis using a polyclonal GlnB antibody, as described above. The deadenylylation rate in WCH15 without IPTG was similar to the rate in the glnB deletion strain (Fig. 3A), confirming the very low expression level of glnB (and glnK) in WCH15 without IPTG. When we increased the GlnB concentration towards wild-type levels (by adding various concentrations of IPTG), the rate of GS–AMP deadenylylation per GS-dodecamer was proportional to the induced P II * concentration (Fig. 3B). Because the variation in the IPTG concen- tration primarily affects glnB expression, this result demonstrates that signal transduction through the GS deadenylylation cascade can be controlled by GlnB (glnK is hardly expressed in medium containing ammo- nia) [32,33,36]. Surprisingly, when the P II * concentra- tion was around and above the wild-type level of 87 ngÆmg )1 protein (open circle in Fig. 3B), the rate of GS–AMP deadenylylation per GS-dodecamer was insensitive to (small) variations in the P II * concentra- tion. Consequently, in wild-type cells, the response coefficient of the GS–AMP deadenylylation rate per GS-dodecamer with respect to P II * concentration was 0. In a narrow region around the wild-type level, P II * concentration did not control deadenylylation rate, although it did when subject to a more sizeable reduc- tion in its concentration (Fig. 3C). If one were to interpret the experimental data (Fig. 3) so as to indicate that, in the P II * concentration range from 0% to 20% below the wild-type level, the GS–AMP deadenylylation rate per GS-dodecamer var- ies linearly with P II * activity, the corresponding response coefficient increased from 0.0 to 0.9 (Fig. 3C). A further increase in the P II * concentration from 20% below the wild-type level to wild-type level, resulted in an abrupt decrease in the response coefficient from 0.9 to 0. Because of the inaccuracy of the measured rates and P II * concentrations we cannot exclude nonlinear variation in the deadenylylation rate when the P II * concentration is below the wild-type level, and hence we cannot be sure about these precise numbers. What- ever the exact kinetics of this variation, it is evident that there is a rather abrupt change in the control of the deadenylylation rate by P II * just below the wild-type P II * concentration. As shown above, the GS–AMP deadenylylation rate per GS-dodecamer was constant around and above the wild-type P II * concentration. The mean value of this constant rate (d[)n] ⁄ dt) is 0.29 s )1 . To determine the extent to which the amount of P II * determines the W. C. van Heeswijk et al. GlnB: ultrasensitive versus subtle control FEBS Journal 276 (2009) 3324–3340 ª 2009 The Authors Journal compilation ª 2009 FEBS 3327 cellular GS–AMP deadenylylation rate, the cellular GS total concentration of these cultures was measured (Fig. 4). At P II * > 50 ngÆmg )1 protein, including the wild-type level, total GS concentration was virtually constant (at $ 4 lgÆmg )1 protein). As a result, the absolute cellular GS–AMP deadenylylation rate was virtually constant around and above the wild-type P II * level (data not shown): the concentration of P II * exerts no control on the absolute cellular GS–AMP deadeny- lylation rate at or above the wild-type P II * level. In cells grown in the absence of ammonia, P II * at wild-type levels does not control the GS–AMP deadenylylation rate either We were also interested in the control exerted by GlnB on the deadenylylation reaction in cells grown in the absence of ammonia. Again we induced GlnB to vari- ous levels by growing the GlnB-tuneable strain WCH15 at various IPTG concentrations overnight in minimal medium with 22 mm glucose, without ammo- nia, but with 14 mml-glutamine (N-poor) [41], to A 600 $ 0.3. At this growth stage, GS was almost com- pletely in the native form and P II * was in the P II *– UMP form (data not shown). After a subsequent 15-min incubation in the presence of 30 mm ammonia, Time (s) GS adenylylation (n) 0 2 4 6 8 10 12 A Molar ratio (P II *) 3 / ATase GS–AMP deadenylylation rate (–n/s) 0.0 0.1 0.2 0.3 0.4 0123 B Response coefficient 0.0 [P II * ] (ng·mg –1 protein) 0 25 50 75 100 125 150 175 [P II * ] (ng·mg –1 protein) 0 25 50 75 100 125 150 175 020 40 60 70 80 100 140 0.2 0.4 0.6 0.8 1.0 C Fig. 3. Control of P II * on the GS–AMP deadenylylation rate per GS-dodecamer in vivo. Cells were grown in the presence of ammo- nia. (A) Deadenylylation of GS–AMP after removal of ammonia at time zero. Open circles, YMC10 (wild-type); open squares, RB9060 (DglnB). The closed symbols depict WCH15 grown in the presence of various concentrations of IPTG (to prevent overcrowding of the figure only some cultures are shown) as follows: circles, 0 l M; squares, 25 l M; triangles, 100 lM; inverted triangles, 300 lM. The curves result from the fitting of the data, as described in Materials and methods. Black lines, YMC10 and RB9060; dotted lines, WCH15. (B) Dependence of the GS–AMP deadenylylation rate per GS-dodecamer on the cellular P II * concentration. The deadenylyla- tion rate was calculated as the rate in the inflection point of the fit- ted curves, shown in (A) (see Materials and methods). The cellular P II * concentration was measured by western blotting. The different points are the mean of three independent cultures of RB9060 (open squares) or WCH15 containing the same IPTG concentration (closed circles); for YMC10 (open circles) five independent cultures were examined. Error bars indicate the SEM. The dotted line is a result of two linear regression fits of the data points. The extra abscissa on top of the figure indicates the molar ratio of P II *-tri- mer ⁄ ATase-monomer. The cellular ATase concentration was mea- sured from YMC10. (C) Response coefficient of the GS–AMP deadenylylation rate per GS-dodecamer with respect to P II *. The response coefficient (R m PIIÃ ) was calculated numerically using the formula described in Materials and methods with the dotted line of (B) as the dataset. Open circle, calculated response coefficient at the P II * concentration of wild-type YMC10. GlnB: ultrasensitive versus subtle control W. C. van Heeswijk et al. 3328 FEBS Journal 276 (2009) 3324–3340 ª 2009 The Authors Journal compilation ª 2009 FEBS GS had been modified towards GS–AMP because of the de-uridylylation of P II *–UMP forming P II *, which stimulates adenylylation. The nitrogen source was then removed, as described above, and samples were taken at the indicated time points to determine the adenyly- lation state of GS (Fig. 5). As shown in Fig. 5B, the GS–AMP deadenylylation rate per GS-dodecamer was now completely indepen- dent of the cellular P II * concentration at an average d[)n] ⁄ dt = 0.12 s )1 . Consequently, the response coeffi- cient of the GS–AMP deadenylylation rate per GS-dodecamer with respect to P II *, as defined above, was 0 and independent of the P II * concentration (data not shown; see Materials and methods for the calcula- tion of these response coefficients). This result indicates that if cells have been pregrown in medium without ammonia, the P II * concentration does not control the GS–AMP deadenylylation rate per GS-dodecamer. At a constant GS–AMP deadenylylation rate per GS-dodecamer, the cellular GS–AMP deadenylylation Time (s) 0 20406080100120140 GS adenylylation (n) 0 2 4 6 8 10 12 A [P II *] (ng·mg –1 protein) 100 150 200 250 300 GS–AMP deadenylylation rate (–n/s) 0.0 0.1 0.2 0.3 0.4 B Fig. 5. In vivo GS–AMP deadenylylation per GS-dodecamer at vari- ous cellular P II * concentrations. Cells grown in the absence of ammonia were incubated with 30 m M ammonia for 15 min. (A) Deadenylylation of GS–AMP after removal of ammonia at time zero. Time is given in s. The adenylylation state of GS is expressed in terms of the average number AMP moieties per GS dodecamer (n). Open circles, strain YMC10 (wild-type); open squares, strain RB9060 (4glnB). Closed symbols depict strain WCH15 grown in the presence of IPTG at various concentrations (to prevent over- crowding of the figure only some cultures are represented) as fol- lows: squares, 25 l M; triangles, 75 lM. The curves result from fitting of the data, as described in Materials and methods. Black lines, YMC10 and RB9060; dotted lines, WCH15. (B) Dependence of the GS–AMP deadenylylation rate (n ⁄ s) on the cellular P II * con- centration. The deadenylylation rate was calculated as the rate in the inflection point of the fitted curves shown in (A) (see Materials and methods). The cellular P II * concentration was measured by western blotting. Each closed circle (WCH15) and open circle (YMC10) is the mean of two experiments (the error bars indicating the standard error of the mean). The closed squares (WCH15) and open square (strain RB9060) are data from single cultures. [P II *] (ng·mg –1 protein) 0 25 50 75 100 125 150 175 [GS total ] (µg·mg –1 protein) 0 2 4 6 8 10 12 Fig. 4. Dependence of the GS total concentration on the cellular P II * concentration. Cells were grown in the presence of ammonia. Cul- tures are the same as in Fig. 3. Cellular concentrations of GS total and P II * were measured by western blot analysis, as described in Materi- als and methods. For the P II * dataset and symbols see Fig. 3B. W. C. van Heeswijk et al. GlnB: ultrasensitive versus subtle control FEBS Journal 276 (2009) 3324–3340 ª 2009 The Authors Journal compilation ª 2009 FEBS 3329 rate might change if the cellular GS total concentration varied with the P II * concentration. The cellular GS total concentration was again measured by western blotting using polyclonal GS antibody. GS total was independent of the P II * concentration (24–30 lgÆmg )1 protein; except for cultures containing 145 ngÆmg )1 protein of P II *, in which a mean GS concentration of $ 8 lgÆmg )1 protein was measured). Therefore, both cellular GS–AMP deadenylylation rate and GS–AMP deadenylylation rate per GS-dodecamer were virtually independent of the P II * concentration (d[GS– AMP] ⁄ dt = 0.24–0.3 mgÆg protein )1 Æs )1 ). This rate was a factor of 2.4 higher than the corresponding rate in cells grown in the presence of ammonia, which agrees with the qualitative increase reported earlier [33]. Cells grown in the absence of ammonia: GlnB versus GlnK Above we measured the primary effect of modulating the expression of glnB in terms of the concentration of P II *, which corresponds to the sum concentration of GlnB and GlnK. The absence of variation in the deadenylylation rate with the increase in GlnB, corre- sponded to an independence of the deadenylylation rate of IPTG and hence expression of the glnB operon, and therefore by definition of GlnB. However, because GlnK might also vary with the induction of more GlnB, this may not necessarily mean an absence of direct control by GlnB on the deadenylylation rate; changes in GlnK might have compensated for the effects of the changes in GlnB. We therefore estimated the concentration of GlnK. [P II *] for wild-type YMC10 grown in the absence of ammonia was 230 ngÆmg )1 protein (Fig. 5B). Because expression of glnB is constitutive [42], the GlnB con- centration in YMC10 in this medium should be 87 ngÆmg )1 protein, as in medium containing ammo- nia. Thus, the ratio [GlnK] ⁄ [P II ] in wild-type cells grown in medium without ammonia should be close to (230)87) ⁄ 87 = 1.7. This is a much smaller ratio than the 500 mentioned as an unpublished observation by Javelle et al. [43]. Because that observation was not documented, the reason for the difference is uncertain. First, the unpublished observation was made in a med- ium with 10-fold lower glutamine concentrations. Sec- ond, the minimal medium was phosphate buffered, whereas in our experiments the medium was buffered with Mops. The phosphate concentration in minimal medium may be relevant because Senior [44] observed a 10-fold increase in GS activity when the phosphate concentration in the minimal medium used was increased 12.5-fold. However, it remains to be seen whether that would be similar for [GlnK]. Third, there was a different strain background (ET8000 versus YMC10; difference in DNA gyrase). Figure 5B proves that the GlnK⁄ GlnB ratio in our wild-type cells, grown in the absence of ammonia, can- not have been 500. If [GlnB] were only 0.2% of [P II *] then reduced glnB expression by the IPTG-induction strategy could never have reduced P II * by > 0.2%. In fact, it was reduced by > 50% in the experiment in which IPTG was absent. Because the ratio [GlnK] ⁄ [GlnB] in wild-type cells grown in the absence of ammonia is only 1.7, if anything, GlnB should repress glnK, and because the primary modulation is that of an increase in the expression of glnB, the increase along the abscissa in Fig. 5 should correspond to the same or a slightly larger increase in [GlnB]. Consequently, neither P II * nor GlnB itself control the deadenylylation rate in cells grown in the absence of ammonia. In cells grown in the absence of ammonia, GlnK is present. [P II *] for RB9060 (glnB-deletion strain) may be equated to [GlnK] (100 ngÆmg )1 protein) (Fig. 5B). Because in this experiment expression of glnB is depen- dent only on [IPTG] and glnK expression is negatively regulated by GlnB, the increase in [P II *] from 100 to almost 300 must imply an increase in [GlnB ] from 0 to 300 or at most 400 ngÆmg )1 protein (the latter if GlnK were to decrease to 0 with increasing [GlnB]). None of this alters the fact that this figure shows that the GS deadenylylation rate does not vary with [GlnB], P II *orglnB gene expression. Hence neither glnB nor GlnB control the deadenylylation rate when cells are pregrown in the absence of ammonia; and nor does the sum of GlnK and GlnB. Therefore, the conclusion of a lack of (ultra)sensitivity in the cascade is not compromised by the fact that the antibody we used to detect GlnB cross-reacts with GlnK. The abrupt change in control by P II * occurs at a P II *-trimer/ATase-monomer molar ratio of 1 The cellular ATase concentration of wild-type strain YMC10, as determined from the two independent cul- tures used in Figs 3 and 5, was 0.18 lgÆmg )1 protein (SEM 7 ngÆmg )1 protein), as measured by western blot analysis using a polyclonal ATase antibody. Expres- sion of the glnE gene, which encodes ATase, is not regulated by the nitrogen status of the cell [42]. This makes it unlikely that the intracellular ATase concen- tration depends on the GlnB or P II * concentration. Assuming that the ATase monomer concentration was 0.18 lgÆmg )1 protein throughout, the molar ratio of GlnB: ultrasensitive versus subtle control W. C. van Heeswijk et al. 3330 FEBS Journal 276 (2009) 3324–3340 ª 2009 The Authors Journal compilation ª 2009 FEBS P II *-trimer to ATase-monomer was calculated for all P II * concentrations [see the extra abscissa (on top) in Fig. 3B]. The rate of GS–AMP deadenylylation per GS-dodecamer changed from being dependent on the P II * concentration to being independent of it around the point at which the molar ratio of the P II *-trimer to ATase was 1. This suggests that above this ratio the ATase is fully saturated with P II *-trimer. Moreover, it suggests that ATase cannot be stimulated by binding more than one P II *-trimer. Additional experimentation should verify this suggestion. P II * uridylylation is competent kinetically It is the uridylylated form of GlnB that stimulates the deadenylylation activity of ATase [22]. Therefore, upon removal of ammonia, the GlnB must be uridylylated before deadenylylation can be set in motion. An expla- nation for the lack of control by wild-type levels of P II * on GS–AMP deadenylylation may be that P II * uridylylation might not keep up with the increase in P II * concentration. UTase may attain its V max at a P II * concentration far below the wild-type concentra- tion of the latter (K m (½P II Ã WT ). To examine this possibility, the uridylylated fraction of P II *, i.e. P II *– UMP ⁄ P II * total , was measured in the same samples used to measure the deadenylylation of WCH15 grown in the presence of ammonia at various P II * induction levels (see above). The two forms of P II * were distin- guished by western blot analysis using a high-resolu- tion tricine gel (see Materials and methods) [33]. The uridylylated fraction (i.e. P II *–UMP ⁄ P II * total ) was determined as described in Materials and methods. As shown in Fig. 6A, at all P II * concentrations, the uridylylation of P II * was almost complete 30 s after the removal of ammonia. The cellular P II * uridylyla- tion rate (not its fractional P II *–UMP ⁄ P II * total uridyly- lation rate) appeared to increase proportionally with P II * concentration (Fig. 6B). Consequently, at P II * levels > 50 ngÆmg )1 protein, the percentage uridylyla- tion at any time after the removal of ammonia was independent of the concentration of P II *, as suggested by Fig. 6A. However, uridylylation of P II * in WCH15 induced with 25 lm IPTG was slower and incomplete compared with cultures induced with higher IPTG concentrations (Fig. 6A). It is possible that at this induced P II * concentration (WCH15 induced with 25 lm IPTG) [P II *] is (far) below the K m of the uridylylation reaction and therefore slower than that in cultures with a (much) higher P II * concentration. This result suggests that the P II * uridylylation reac- tion per se is quick enough for P II *–UMP to activate deadenylylation. The reaction may still progress but only because of a progressing change in the signals impinging on uridylyl transferase (such as glutamine or 2-oxoglutarate). Time (s) P II *-UMP / P II * total 0.0 0.2 0.4 0.6 0.8 1.0 A [P II *] (ng·mg –1 protein) 0 20406080100 0 25 50 75 100 125 150 175 P II * uridylylation rate (ng·mg –1 protein·s –1 ) 0 1 2 3 4 B Fig. 6. Uridylylation of P II * in vivo. Cells were grown in the pres- ence of ammonia. Cultures are the same as in Fig. 3. (A) Uridylyla- tion of P II * after removal of ammonia at time zero. Open circles, YMC10 (wild-type). The closed symbols depict WCH15 grown at var- ious concentrations of IPTG as follows: circles, 25 l M; squares, 75 l M; triangles, 150 lM; inverted triangles, 300 lM. The curves result from fitting of the data as described in Materials and methods. Black line, YMC10; dotted lines, WCH15. (B) Dependence of the P II * uridylylation rate on the cellular P II * concentration. The uridylylation rate was calculated as the initial rate of the fitted curves shown in (A) (see Materials and methods). The cellular P II * concentration was measured by western blotting. The different points are the means of two or three independent cultures and correspond to those in Fig. 3. Error bars indicate the standard error of the mean. The line results from a linear regression calculation of the data points. W. C. van Heeswijk et al. GlnB: ultrasensitive versus subtle control FEBS Journal 276 (2009) 3324–3340 ª 2009 The Authors Journal compilation ª 2009 FEBS 3331 As to the initial uridylylation rate, the response coefficient with respect to the P II * concentration was close to 1 for the P II * concentration range examined (Fig. 6B; detailed analysis not shown). Apparently, the uridylylation reaction appears to be noncoopera- tive with respect to P II *, in agreement with in vitro data [45]. Figure 7 directly compares the transient uridyly- lation of P II * (i.e. a decrease in P II * ⁄ P II * total ) with the transient deadenylylation of GS–AMP per GS-dode- camer (GS–AMP ⁄ GS total ,adecrease in the adeayly- lation state of GS) at various engineered P II * concentrations. We conclude that the former (Fig. 7, open squares) preceded the latter, but with increasing cellular P II * concentration, the rate of uridylylation approached the rate of deadenylylation. P II * has no control over the steady-state GS adenylylation state in the presence of ammonia As described above, to study the deadenylylation rate as a function of induced GlnB, strain WCH15 was grown overnight in the presence ammonia and at vari- ous concentrations of IPTG to A 600 $ 0.3 before the ammonia was removed. Consequently, the GS adenyly- lation state (which we denote by n) before the ammo- nia is removed (at time zero in Fig. 3A), represents the steady-state adenylylation state of these cells in the presence of ammonia. As shown in Fig. 8 (left), varia- tion in the P II * concentration around the wild-type level did not change this steady GS adenylylation state. Consequently, at wild-type levels of P II *, the response coefficient of the steady-state GS adenylylation state towards P II * concentration was 0 : P II * did not control the steady-state GS adenylylation state. At cellular P II * concentrations < 40 ngÆmg )1 pro- tein, the steady-state GS adenylylation states appeared slightly higher than at higher P II * concentrations. The difference in the adenylylation state of the glnB dele- tion strain ($ 11) compared with that of the wild-type strain ($ 9) should correspond to a decrease in active (unmodified) GS by $ 60%, if the total GS concentra- tion was the same in both strains. With the total GS concentration (Fig. 4), one can calculate the cellular (active) GS concentration as function of the cellular P II *. Perhaps surprisingly, the cellular nonadenylylated GS concentration was approximately constant over the range of P II * measured, and also at low P II * concen- trations (see dotted line in Fig. 8, right). The slight increase in adenylylation state and the increase in GS total concentration at low P II * concentrations appear to compensate for one another, perhaps reflecting homeostatic regulation. Discussion In this study, we tested quantitatively in vivo and under two relevant growth conditions, whether signal transduction from ammonia depletion to GS–AMP deadenylylation is highly sensitive to the concentration of the pivot of the GS cascade, i.e. GlnB. It was not. In fact it was not sensitive at all to the concentration of GlnB (P II *) around the wild-type level of the latter. Neither the steady-state extent of adenylylation of GS, nor the rate at which GS–AMP became deadenylylated upon ammonia deprivation, depended on glnB gene expression (as modulated by IPTG) or on the concen- tration of P II * (i.e. GlnB–UMP plus GlnK–UMP). This most direct in vivo test refutes a signal-amplifica- tion function proposed for this cascade in vivo under Time (s) 0 20 40 60 80 100 120 140 GS–AMP/GS total 0.0 0.2 0.4 0.6 0.8 1.0 P II */P II * total 0.0 0.2 0.4 0.6 0.8 1.0 Fig. 7. Comparison between the transient uridylylation of P II * and the transient deadenylylation of GS–AMP. See also Figs 3 and 6. Cells had been grown in the presence of ammonia. Uridylylation of P II * and deadenylylation of GS–AMP were measured after removal of ammonia at time zero. The data points are raw data; the curves are connections between the data points and do not result from fitting of the data. Transient deadenylylation reactions at different P II * concentrations: open circles, YMC10. The closed symbols depict WCH15 at various concentrations of IPTG as follows: cir- cles, 0 l M; squares, 25 l M; triangle, 300 lM. Only one transient uridylylation of the fractional P II * ⁄ P II * total is shown because the transient uridylylation of the fractional P II * ⁄ P II * total after removal of ammonia was independent of the P II * concentration (Fig. 6). Open squares, wild-type YMC10. To simplify the comparison, the tran- sient deadenylylation of GS–AMP is shown as a decrease of the fractional adenylylation level (left y-abscissa), and the transient uridylylation of P II * as a decrease of the fractional native P II * level (right y-abscissa). Both were calculated from data of Figs 3 and 6, respectively. GlnB: ultrasensitive versus subtle control W. C. van Heeswijk et al. 3332 FEBS Journal 276 (2009) 3324–3340 ª 2009 The Authors Journal compilation ª 2009 FEBS at least two important physiological conditions. Because this cascade often figures as a model for signal transduction, this conclusion should be important for understanding signal transduction more generally. In view of the possible role of the paralogue of GlnB, GlnK, we tested for signal amplification in two times two ways: (a) we determined the sensitivity towards variation in expression of the glnB gene around its wild-type level, and (b) we determined how strongly function varied with the level of GlnB plus GlnK in the same experiment. We performed this experiment first under a condition in which we con- firmed that GlnK was virtually zero, and then under a condition in which, at the zero level of GlnB, GlnK was substantial. Because both covariations were zero in both experiments, this implies that at physiological levels of GlnB, the dependence of function on GlnB is zero, independent of whether one takes any possible variation of GlnK into account. It is important that we emphasize here that we discuss dependence in terms of the effect of small variations in GlnB around its wild-type level. For larger variations, the issue is more complex, but ultrasensitivity was still not observed (Fig. 3 and below). The mechanistic explanation for this outcome may well be that the models leading to the prediction of zero-order ultrasensitivity [3,4] do not apply in vivo [8,9], that the in vivo kinetics and abundances were such that they did not give rise to zero-order ultrasen- sitivity, or that gene expression mediated adaptation involving GlnK or metabolic adaptations prevented kinetic scenarios from being enacted. As to the first possibility, the GS adenylylation cascade differs from the cascades modelled in these studies: the two reac- tions catalysed by ATase are activated by different activators, GlnB and GlnB–UMP (Fig. 1), which are in balance at steady-state growth. The third explana- tion is unlikely because we determined GS deadenyly- lation as a function of the sum concentration of GlnB and GlnK (P II *) and of the induction of glnB expres- sion. With respect to the metabolic adaptations, that of the concentration of 2-oxoglutarate is also unlikely. Under nitrogen-limiting conditions, [P II *] changes by $ 5 lm, whereas under nitrogen excess, [P II *] varied from 0 to 3 lm. This variation is much less than the reported intracellular concentration of 2-oxoglutarate under those conditions (from $ 0.1 to $ 0.9 mm) [44]. Our observation of a steady-state GS adenylylation level being independent of [GlnB] around its wild-type level (Fig. 8) is in agreement with a computer simula- tion of the GS adenylylation bicyclic cascade by Muta- lik et al. [46]. Our observations leave us with the puzzle of a func- tional explanation for the existence of GlnB: why should control by P II * be absent altogether, and what then is the function of the cascade and of its pivot GlnB? The problem is reinforced by the observation that, in cells grown in the absence of ammonia, dele- tion of GlnB hardly affected the deadenylylation rate of GS–AMP. The pivotal protein GlnB appeared to be redundant. One functional explanation for redundant pheno- types is that of the conditional phenotype, i.e. some proteins only function under special conditions [47]. [P II *] (ng·mg –1 protein) [P II *] (ng·mg –1 protein) Non-adenylylated GS (µg·mg –1 protein) 0 1 2 3 0 25 50 75 100 125 150 175 0 25 50 75 100 125 150 175 GS adenylylation state (n) 0 2 4 6 8 10 12 Fig. 8. Dependence of the steady-state adenylylation state and dependence of the concentration of nonadenylylated GS on the P II * concen- tration. Cells were grown in the presence of ammonia. Cultures were the same as in Fig. 3. (Left) Steady-state adenylylation state at various cellular P II * concentrations (see time zero of Fig. 3A). For the P II * dataset and symbols see Fig. 3B. The dotted line is not a result of fitting of the data points. (Right) Calculated concentration of nonadenylylated GS (from Fig. 8, left and Fig. 4) at various cellular P II * concentrations. Symbols are as in Fig. 8 left. The dotted line (see text) has been drawn by hand. W. C. van Heeswijk et al. GlnB: ultrasensitive versus subtle control FEBS Journal 276 (2009) 3324–3340 ª 2009 The Authors Journal compilation ª 2009 FEBS 3333 [...]... functions and by using the described fitting function 20 or 30 s after the removal of ammonia rather than for the in ection point of the curve This did not change Figs 3B and 5B qualitatively Another consequence of GlnB UMP production during the deadenylylation reaction is that the variable at the abscissa of Figs 3B and 5B cannot quite be taken to represent PII*–UMP (instead of induced PII*), even though the. .. function (and phenotype) The question arises as to what could be the mechanism of the subtle regulation by and around GlnB Here, a key observation may be the correlation of the expression of the GlnB paralogue GlnK with the change of control by GlnB on the deadenylylation rate When cells are grown in the absence of ample ammonia, glnK is expressed [33,36], and heterotrimers can be formed when both GlnB, and. .. phenotype differed: in the former case, large reductions in the expression level of glnB did affect the GS deadenylylation rate and a complete knockout of GlnB did do so very strongly [33], whereas in the latter case there was complete independence of PII* (b) The effect of reducing the concentrations of P*II in cells grown in the presence of ammonia depended on the magnitude of the reduction, being zero for... previously [33] Scanning of the autoradiograms and the determination of intensity of the PII* and PII*–UMP bands were as described above Some gels did not result in a complete separation of the uridylylated and native PII* forms In that case, the integrated density of the two bands was fitted with an equation containing two Gaussian functions using the computer program sigmaplot (Jandel Scientific) ð3Þ... present [49,54] In vitro, uridylylated GlnK ⁄ GlnB heterotrimers can stimulate the deadenylylation of GS–AMP This may explain the absence of a GlnB phenotype in cells grown in the absence of ammonia The GlnK could already suffice to saturate ATase, modulation of the GlnB would then have no further effect, and hence GlnB expression and P*II would have no control In cells grown in the presence of ammonia,... cloning the NotI fragment of pWVH93, containing the promoter cassette, into the EcoNI site of pWVH90, after both sites had been blunted with the Klenow fragment of DNA polymerase I The endogenous ribosomal binding site of the glnB gene resides downstream the EcoNI site The endogenous promoters of glnB [42,64,65] are still present on the chromosome after homologous recombination of the DNA between the. .. [59] and ligated into pFC13 digested with BamHI and SspI, resulting in pWVH112; and (b) inserting a linker containing the restriction sites SfiI, NotI and BssHII with on both ends sticky BamHI sites (5¢-GATCCGCGCGCGGCCGCCTAGGCC G-3¢) into the BamHI site of pWVH112, resulting in pWVH116 The glnB gene with the promoter cassette upstream, was isolated as a SfiI fragment of pWVH102 and ligated into 3336 the. .. Jensen PR, Westerhoff HV & Michelsen O (1993) Excess capacity of H+-ATPase and inverse respiratory control in Escherichia coli EMBO J 12, 1277–1282 Kahn D & Westerhoff HV (1991) Control theory of regulatory cascades J Theor Biol 153, 255–285 Ninfa AJ, Jiang P, Atkinson MR & Peliska JA (2000) Integration of antagonistic signals in the regulation of nitrogen assimilation in Escherichia coli Curr Top Cell... heterotrimers To further substantiate of these mechanistic explanations, more research will be required Such research should also clarify the minor uncertainties left in this study As described above, the concentration of GlnB was measured as PII*, i.e as a sum of GlnB and GlnK, including the uridylylated form of both proteins Cells grown in the presence of ammonia barely express glnK [33,36] Therefore, the measured... same for different induced levels of GlnB for cells grown in the absence of ammonia, when GlnK is expressed In either case, the uncertainty in the ratio GlnB ⁄ PII* affects our conclusion that neither GlnB nor GlnB and GlnK combined (PII*) control deadenylylation: the observed response coefficient to IPTG was 0 and when glnK expression was not activated by excess ammonia, independent of the PII* concentration . The pivotal regulator GlnB of Escherichia coli is engaged in subtle and context-dependent control Wally C. van Heeswijk 1 ,. chain. The functional activity of the protein at the bottom of the hierarchy depends on the modification state of that protein. The advantage of modulating